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Chapter 4

Biochemical and Genetic Mechanisms of Insecticide Resistance

Downloaded by UNIV OF MISSOURI COLUMBIA on April 12, 2017 | http://pubs.acs.org Publication Date: February 23, 1990 | doi: 10.1021/bk-1990-0421.ch004

Thomas M. Brown Department of Entomology, Clemson University, Clemson, SC 29634

The biochemical mechanisms of insecticide resistance are categorized as affecting insecticide pharmacokinetics or pharmacodynamics. The enzymes and targets involved are reviewed from the perspective of understanding the genetic basis for each mechanism. This perspective is important since insecticide resistance is a problem in population genetics and each case of resistance must be managed according to the particular resistance gene, or combination of genes, present. The source of the insecticide resistance problem is traced ultimately from biochemical mechanisms to changes in nucleic acid chemistry of genes which confer resistance. Advances in genetics and recombinant DNA have provided the opportunity for detailed study of genes that produce biochemically resistant populations. Insecticide Resistance Mechanisms. These can be catagorized as behavioral avoidance and physiological changes which allow survival upon contact with the insecticide. Major physiological mechanisms are categorized by whether they influence insecticide pharmacokinetics (the penetration, distribution, metabolism, and elimination of a drug or other xenobiotic) or insecticide pharmacodynamics (the interaction of a drug or other xenobiotic with its site of action). Consideration will be given to the recent advances in the molecular genetics of resistance mechanisms known or suspected to be important (Table I). This review will focus upon detoxication as a pharmacodynamic mechanism and also pharmacodynamic mechanisms in which various targets of insecticides, often proteins in the nervous system, become less sensitive to poisoning. Besides detoxication, another pharmacokinetic mechanism of resistance, retarded penetration, has been identified and mapped to linkage group III of the house fly where it is known as Pen &) or tin (2). This gene imparts low resistance; however, its combination with other mechanisms can be synergistic in producing higher than expected resistance. Because the biochemical basis of this mechanism is unknown, it will not be discussed further in this paper. The genetic basis of insecticide resistance is not well understood. Many genes for resistance traits have been mapped to chromosomes of the house fly, Musca domestica (3 4) Drosophila melanogaster (78), and mosquitoes, Aedes aeqypti and Culex quinquefasciatus (5 6). In some cases, the expression of the biochemical f

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0097-6156/90/0421-0061$06.00/0 © 1990 American Chemical Society

Green et al.; Managing Resistance to Agrochemicals ACS Symposium Series; American Chemical Society: Washington, DC, 1990.

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MANAGING RESISTANCE TO AGROCHEMICALS

mechanism, such as increased activity of enzyme or decreased sensitivity of target, has also been mapped to the same locus as resistance; however, few studies have established the genetic mechanism responsible for resistance by mapping the biochemical polymorphism or analyzing the gene sequence.


Table I. Principal Physiological Mechanisms of Insecticide Resistance Pharmacokinetic mechanisms

Pharmacodynamic mechanisms (reduced sensitivity of target)

Decreased penetration

Acetylcholinesterase Sodium ion channel y-Aminobutyric acid (GABA) receptor

Enhanced detoxication Monooxygenase Arylester hydrolase Carboxylester hydrolase Catalytic hydrolysis Insecticide sequestering Glutathione S-transferase

Beyond these examples, there is very little known of the genetics of insecticide resistance in many important insects of agriculture which are the targets for much of the insecticide used in the world; e. g., boll weevil, tobacco budworm, codling moth, and Colorado potato beetle are practically unknown genetically. The present challenge is to apply new techniques of molecular genetics to gain a better understanding of resistance in many of the pests of greatest economic impact for the future development of insecticide resistance management. A case in point is the unraveling of resistance to methyl parathion in the tobacco budworm, Heliothis virescens, which is a major pest of cotton as well as tobacco. In South Carolina, there is very severe, stable resistance. Although pyrethroid insecticides are very effective and there is no resistance to them in South Carolina at this time, it would be very useful to understand the genetic basis of methyl parathion resistance in case resistance to pyrethroids should arise in the future or spread eastward from Texas where it has been detected. Recent investigations with this pest will be described to illustrate certain mechanisms. Enzymes Catalyzing Insecticide Biotransformation Even a rather simple insecticide such as methyl parathion is transformed by insects in a complex manner. The parent insecticide is activated to methyl paraoxon, which is a more potent inhibitor of the target, acetylcholinesterase in the nerve (Figure 1). This activating desulfuration is catalyzed by monooxygenases. Both the parent and the oxon are subject to detoxication by monooxygenase and glutathione transferase, while the oxon is also more labile to hydrolysis. Monooxygenases. These enzymes are important in the detoxication of pyrethroid, carbamate, organophosphorus and other classes of insecticides CD. They are microsomal, membrane associated enzymes which catalyze reactions in which one atom from molecular oxygen is inserted into the insecticide and the second oxygen atom is reduced to form water. Catalysis depends on the close association of the hemecontaining cytochrome P450 terminal oxidase with NADPH cytochrome C reductase for electron transport, and it also depends on availability of NADPH and oxygen. Monooxygenase Inhibitors as Synergists. Inhibitors of cytochrome P450 are used as preliminary diagnostic synergists (&) to detect resistance based on a mechanism of


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Biochemical & Genetic Mechanisms ofInsecticide Resistance

monooxygenase catalyzed detoxication (Figure 2). Fenvalerate resistance in diamondback moth in Taiwan and also in cotton bollworm, Heliothis armigera, in Australia can be synergized with piperonyl butoxide (9 10). In permethrin-resistant Heliothis virescens, the tobacco budworm, piperonyl butoxide was not synergistic; however, l,2,4-trichloro-3-(2-propynyloxy)benzene (Figure 2) was synergistic with permethrin (H). In resistant Egyptian cotton leafworms, Spodoptera littoralis , propynyl monooxygenase synergists were 2.5-fold more active than piperonyl butoxide when used with monocrotophos (12). Lack of synergism must be interpreted cautiously, especially with insecticides which are metabolically activated. Bioactivation of organophosphorothioate, formamidine and certain cyclodiene insecticides is catalyzed by monooxygenases. Piperonyl butoxide was synergistic with the carbamate, propoxur; however, in the same strains of house flies, it was antagonistic with the phosphorothioate, diazinon, which requires monooxygenase desulfuration to form diazoxon, the oxon inhibitor of acetylcholinesterase (12). Chordimeform was also antagonized by piperonyl butoxide in the Australian cattle tick (14). Chordimeform is bioactivated to JV-dealkylation products which are much more potent octopamine mimics in the fire fly (15). In methyl parathion resistant tobacco budworms, larvae were treated, lots of 10 were homogenized, and methyl parathion was recovered by solid phase extraction and analyzed by reversed phase high performance chromatography with ultraviolet detection. Unexpectedly, the resistant strains lost methyl parathion at a lesser rate than the susceptible Florence 1987 strain (Figure 3). Independently, T. Konno et. al. have found a slower bioactivation of [ C]-methyl parathion in another resistant strain of this pest (22). We observed that both l,2,4-trichloro-3-(2-propynyloxy)benzene and S,S,Stributylphosphorotrithioate (a hydrolase inhibitor) were synergists for methyl parathion in Woodrow 1983-resistant 35 mg larvae. Topical doses were applied in acetone for a 48 h exposure (Table II). This suggests both monooxygenase and hydrolase involvement in resistance. Others (Konno, T.; Dauterman, W., North Carolina State University, personal communication, 1988) have confirmed our observation of synergism using their NC-86 resistant larvae. They also found the propynyl synergist was more effective with methyl paraoxon (Table II). Both laboratories found less than 2-fold synergism by piperonyl butoxide in methyl parathion resistant strains (16).


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Table II. Synergism of Methyl Parathion in Heliothis virescens, the Tobacco Budworm Strain

Insecticide

NC-S NC-R NC-S NC-R SC-R SC-R

m. paraoxon m. paraoxon m. parathion m.parathion m. parathion m. parathion m. parathion m. parathion

SC-R SC-S

Synergist TCPB

a

TCPBa TCPBa TCPBa none TCPB TBPT TBPT

b b

C

Median lethal dose, mg/kg Insecticide Insecticide + Synergist 5.00 182 10.8 618 2600 28% at 400 28% at 400 815

Ratio

4.74 76.4 12.7 433

1.05 2.38 0.85 1.43

75% at 400 97% at 400 155

2.68 3.46 5.26

Synergist l,2,4-trichloro-3-(2-propynyloxy)benzene at 2857 mg/kg; data from T. Konno and W. Dauterman (see text above). Synergist l,2,4-trichloro-3-(2-propynyloxy)benzene at 2000 mg/kg. Synergist S,S,S -tributylphosphorotrithioate (16).

Genetic Control of Monooxygenases. There are many forms of P450, even within a species, with differences in spectra, substrate specificity, electrophoretic mobilities and


6

64

MANAGING

C r ^ O ^ J - O - ^ ^ - N O ,

RESISTANCE TO AGROCHEMICALS

C^O-J-O-^^-NO,

•

monooxygenase methyl

parathion

methyl paraoxon


Bioactivation

M

s

CH

/

o\ /

S \

/

G

M

M

AH

\

G

M

Detoxication

Figure 1.

Bioactivation and detoxication of methyl parathion. A H is arylester hydrolase, C H is carboxylester hydrolase, G is glutathione transferase, and M is monooxygenase.

piperonyl

butoxide

CH^CBjCHjS—SCr^CH>CHjCH> icr^CH^CHjCHj

T B PT

Figure 2.

CI-

X

^

K

^

/

'

/

h\

chlorfenethol

Diagnostic synergists for insecticide resistance described in the text.


Biochemical & Genetic Mechanisms of Insecticide Resistance


4. BROWN

Figure 3.

Methyl parathion remaining after topical application to tobacco budworm larvae in vivo. Florence 1987 strain was resistant; other strains were susceptible.



66


specificity of inducing agents. Cloning of about 70 P450 genes, primarily from mammals, has provided a systematic classification of these forms based on the amino acid sequences inferred from the nucleotide sequences. There are at least 10 gene families, each having no more than 36% similarity in amino acid sequence to the others, and families II and XI are comprised of subfamilies containing genes coding proteins with no less than 67% similarity to each other (17). Recently, a P450 clone was prepared from phenobarbital induced, diazinon resistant, house flies (li£). While the amino acid sequence near the heme-binding site is conserved compared to others, the house fly gene was assigned a new family, P450VI, because its overall amino acid sequence was insufficiently similar to any previous family. Several forms of P450 are known to exist in the house fly (12.20). Monooxygenase activity is strongly inducible in both insects (21) and mammals (17). The quantity of the enzyme increases 10 to 100-fold upon prolonged exposure to an inducing agent, probably due to a more rapid rate of gene transcription. There is a wide chemical diversity of inducers (Figure 4) which are usually selective for the family of P450 which is induced (12). The relationship of induction to insecticide resistance has not been established. In mice, induction of P450I results from increased transcription of the structural gene as controlled by a cytosolic receptor which is the protein product of the Ah locus (12). Resistance due to monooxygenases is influence by genes ox and md on chromosomes 2 and 5, respectively, in the house fly (2). Similarly, two genes confer monooxygenasebased resistance in Drosophila, and there is evidence that one is regulatory in nature (22). A working hypothesis is that unlinked regulatory and structural gene mutations contribute; however, multiple forms of insect P450 enzymes exist so there could be mutant structural genes on separate chromosomes. Hydrolases. Hydrolytic mechanisms are also important in insecticide resistance, despite the apparent low activities in resistant insects when compared to mammalian enzymes (Table III). Some strains of resistant mosquitoes (23), Tribolium beetles (24), and Indianmeal moth (25) have specific resistance for malathion and similar carboxylester insecticides. This is due to increased catalytic hydrolysis, possibly through production of a more efficient enzyme (2526). Californian tobacco budworms with low level permethrin resistance exhibited twice the normal activity of transpermethrin carboxylester hydrolase (27). It is clear that the insecticide of interest must be used to assay for this type of resistance since 1-naphthyl acetate, a general substrate used to stain for many hydrolases, did not detect malathion carboxylester hydrolase (assayed as the hydrolysis of [ C]-malathion) in starch gel electrophoresis of Culex tarsalis (22). Similarly, fra/ts-permethrin hydrolyzing activity of soybean looper was resolved from 1-naphthyl acetate hydrolyzing activity by polyacrylamide gel electrophoresis (2&). Malathion carboxylester hydrolases were not correlated with activity toward 1naphthyl acetate in Drosophila, Anopheles, and Indianmeal moth (25.29.30). Malathion carboxylester hydrolase activity was genetically linked to chromosome 2 of the house fly as demonstrated in the Wakamatsu strain (21). Malathion resistance in this strain was nearly completely dominant, and the resistance was also linked to chromosome 2. It was found that this highly resistant strain also possessed enhanced glutathione dependent malathion detoxication, as well as acetylcholinesterase which was 76-fold less sensitive to malaoxon. These flies were cross-resistant to the non-carboxylester organophosphorus insecticides fenitrothion and trichlorfon, probably due to glutathione transferase or acetylcholinesterase factors; however, they were fully susceptible to pyrethroid insecticides containing carboxylesters, suggesting that the carboxylester hydrolase was selective. While resistance is highly dominant, malathion hydrolysis in hybrid progeny of Wakamatsu and susceptible house flies was only 60% as much as malathion hydrolysis observed in the homozygous resistant flies. r

14




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Figure 4.

Inducers of monooxygenases for various P450 families or for insect P450.


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68

MANAGING RESISTANCE

TO AGROCHEMICALS

Malathion specific resistance was also fully dominant in Indianmeal moth, while enzyme activity was semidominant, with hybrid offspring having one-half the activity of resistant homozygotes (253- These data in house flies and moths are consistant with a mechanism of catalytic hydrolysis with one resistance allele giving sufficient enzyme activity for survival. Table III. Insecticide Hydrolysis by Enzymes of Mammals and Resistant Arthropods Insecticide Enzyme or preparation


Paraoxon

Activity (nmol/min/mg protein)

House fly carboxylester hydrolase Green peach aphid carboxylester hydrolase Bovine serum albumin Rabbit serum arylester hydrolase Sheep serum arylester hydrolase

Malathion

0.00733 0.256* 0.044 642 1003

Spider mite carboxylester hydrolase House fly carboxylester hydrolase Indianmealmoth midgut preparation Rat serum Rabbit liver carboxylester hydrolase Rat liver carboxylester hydrolase

frans-Permethrin Southern armyworm cuticle preparation* Soybean looper midgut preparation* 1

1

Rat liver carboxylester hydrolase Porcine liver carboxylester hydrolase

(ID (24) (22) (42) (4a)

0.0203 0.937 3.94

(25)

84.8* 4160* 7630; 16700a

(24) (25)

3.62a 23.9a 70.0 322

(2D (22)

(23) (22) (2a)

(23) (2fi)

Vmax

Insects were not reported to be resistant. Amplification of a Normal Hydrolase Results in Insecticide Sequestration. Gene amplification to produce geometric increases in carboxylester hydrolase quantity provides a second type of hydrolytic mechanism in certain resistant insects. This genetic mechanism has been found using radiolabeled cDNA of the gene to estimate the quantity of the gene and its transcription in Culex quinquefasciatus (32) and in Myzus persicae (22)- Resistance is conferred to many organophosphorus insecticides, including non-carboxylesters such as parathion, and to carbamates and pyrethroid in the aphids. The tremendous quantity of the enzyme is a sink for organophosphorus oxon bioactivation products which phosphorylate this enzyme more rapidly than the target. This enzyme, purified from aphids (24), was very slow in hydrolyzing paraoxon compared to mammalian arylester hydrolases (Table III); however, it was twice as fast in recovery from paraoxon inhibition compared to a porcine carboxylester hydrolase (25) and >300 times faster than monomeric carboxylester hydrolase of rabbit liver (2£). No qualitative differences were found in the enzyme, E4, isolated from resistant and susceptible aphids; E4 was one of seven electrophoretic forms of hydrolases observed (24). Recovery indicates that resistance is due to both reaction with the insecticide and a very slow turnover, or catalysis. A similar mechanism of was observed with paraoxon in resistant green rice leafhoppers (37).


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Resistant aphids are identified individually in microtiter plate wells by immunological binding of E4 followed by colorimetric assay with 1-naphthyl butyrate (M). In this aphid, there appears to be no other major resistance mechanism; however, a very similar mechanism in mosquitoes can be accompanied by other mechanisms, including insensitive acetylcholinesterase (3940). Carboxylester hydrolase is strongly inhibited by many organophosphates such as paraoxon. This is the reason that malathion resistance is overcome by its mixture with many other organophosphorus insecticides. In fact, malaoxon is both a substrate and an inhibitor depending on the orientation of the molecule, which contains both carboxylester and phosphate ester chemistry (41). S,S,S-tributylphosphorotrithioate (Figure 2) and triphenyl phosphate, which are not insecticidal, have been used as diagnostic synergists for this mechanism. Because resistance to malathion is increasing, it may become practical to seek synergistic mixtures against resistant pests; however, such mixtures could have much greater toxicity to mammals as well.


f

Arvlester Hydrolase and Parathion Hydrolase. Arylester hydrolase in mammalian serum (4243) catalyzes very efficient hydrolysis of the organophosphorus oxons such as paraoxon (Table III). This enzyme does not hydrolyze the parent phosphorothioate insecticides such as methyl parathion, but only the oxon metabolites such as methyl paraoxon (44). This enzyme is has been referred to as phosphotriesterase; however, a triester is not required as seen in the rapid hydrolysis of 10 organophosphinates by rabbit serum arylester hydrolase (44). This mechanism appears only rarely in birds (453 and appears to be lacking in most insects. Recently, an arylester hydrolase requiring cobalt was isolated from a methyl parathion resistant strain of tobacco budworm (79). A parathion hydrolase has been described to hydrolyze both phosphorothioates and phosphates and its gene, opd, cloned from a plasmid carried by Psudomonas diminuta (4£). Fortunately, insects apparently lack this type of enzyme. Forty years of effective applications of organophosphorus insecticides can be attributed to some extent to the general lack of significant arylester hydrolase activities in pest insects. r

Glutathione -Transferases. These enzymes have been associated with resistance in house flies and in mites, but have not been associated with resistance in as many species as monooxygenases or hydrolases (41). This may be due in part to less testing for this mechanism. In the house fly, resistance due to glutathione S-transferase is linked to chromosome 2 and the mechanism apparently results from a qualitative change in the enzyme (48). Cloning and analysis of glutathione S-transferase genes of mammals and plants has revealed that there are various gene families; however, this analysis has not been accomplished in insects (42). These enzymes are inducible, but generally not to the degree of the monooxygenases (4850). DDT-dehydrochlorinase is a glutathione-dependent enzyme which is inhibited by chlorfenethol (Figure 2). It is linked to chromosome 2 of the house fly (4) • r

Altered Targets

Acetylcholinesterase. Altered acetylcholinesterase less sensitive to organophosphorus and carbamate insecticides has been observed in a wide variety of insects and mites (51). Acetylcholinesterase inhibiting insecticides phosphorylate or carbamylate the serine residue in the active site of the enzyme preventing vital catalysis of acetylcholine. Resistance due to reduced sensitivity to inhibition of this target enzyme has been found in house fly, mosquitoes, green rice leafhopper, and both phytophagous and predacious species of mites. In the house fly, at least two resistance alleles occur. They can be discriminated



70

MANAGING RESISTANCE

TO AGROCHEMICALS

by relative insensitivities to azamethiphos and dichlorvos (52). A Danish strain, 49R, was less sensitive to azamethiphos. Conversely, a German strain, Weymann, was sensitive to azamethiphos, but insensitive to dichlorvos. Individualfliesof these two strains, a susceptible strain and the three possible hybrids could be identified by inhibition kinetics measured in a microtitre plate reader; apparently, both resistance alleles were incompletely dominant. Previous investigations with tetrachlorvinphos resistant house flies indicated that resistance was due to a reduced affinity for binding the insecticide (53). In resistant green rice leafhopper, an acetylcholinesterase with reduced affinity for iV-methyl carbamate insecticides had increased sensitivity to inhibition by longer iV-alkyl groups; however, the inverse relationship was observed against the susceptible enzyme (M). The optimal substitution appeared to be iV-C/i-propyl) and the use of this chemistry in a resistance breaking strategy was discussed. Unfortunately, N-(npropyl)propoxur was inhibitory of neither susceptible nor resistant acetylcholinesterase of predatory mites, Amblyseius potentillae so that this type of vulnerability is not common to all resistant acetylcholinesterases (55). In general, resistant acetylcholinesterases are less sensitive as indicated by a smaller bimolecular reaction constant, ki, for phosphorylation of the active site. In our studies of methyl parathion resistant tobacco budworm larvae, lots of ten larval nervous systems were homogenized and ki was determined We observed 25-fold less sensitivity to inhibition to methyl paraoxon in the resistant strain (Table IV). Table IV. Reduced Rate of Acetylcholinesterase Phosphorylation in Resistant Tobacco Budworm Larvae, Heliothis virescens Strain

Median Lethal Dose, mg/kg methyl paraoxon

ki, M-imin-l ethyl paraoxon

Florence 87 (S) Woodrow 83 (R)

o CO CD

Resistant Hybrid Susceptible


CC

15

Time, h Figure 5.

Spontaneous reactivation of acetylcholinesterase from tobacco budworm larvae.

O

acetylcholinesterase

Recovery Figure 6.

and aging

Comparison of phosphorylated and phosphinylated acetylcholinesterase recovery.


7


72

MANAGING

RESISTANCE TO AGROCHEMICALS

mechanism in a strain of the tobacco budworm in which segregation of high permethrin resistance was demonstrated (61). Other ion channels functioning as neuroreceptors are known targets of insecticides (Figure 7). These include the GABA receptor which is a target of pyrethroids, cyclodienes and avermectin, and the acetylcholine receptor which is the target of nicotine. These receptors are similar in forming a transmembrane pore. Only one sodium ion channel subunit is required for expression in Xenopus oocytes injected with mRNA from its cloned DNA (62): this gene has four similar domains, each containing six transmembrane helices. A putative sodium channel has been cloned from Drosophila and its sequence determined (02); however, the relationship of sequences to resistance is not known. GABA receptor, a target for several classes of insecticides, consists of two subunits, each with four transmembrane helices as determined from cloning and sequence analysis (M). Expression in Xenopus oocytes requires mRNAfromonly one of the subunits (£5). Dieldrin resistant cockroaches were found to have reduced specific binding of picrotoxinin, which acts on the GABA receptor chloride ion channel (££)• The acetylcholine receptor is also a transmembrane ion channel, and it is composed of four different subunits (Q2). Chimeric mRNA from recombinant clones of different species has been expressed in Xenopus oocytes. This is the target of nicotine, a classical agonist for discriminating among receptor subtypes. This receptor has not been evaluated as a resistance mechanism, although it is involved in the process of poisoning by many insecticides. Prospects and Imnerative Research Genetic studies have raised many new questions. In pursuit of answers to these questions, the knowledge gained should lead to improved strategies for resistance management. In conclusion, some of these questions will be considered, and then prospects for genetic research on major pest species will be discussed. Questions Involving Genetic Mechanisms. Gene amplification, the presence of multiple copies of the gene of interest, provides two resistant species with greatly elevated carboxylester hydrolase (see above). While gene amplification is the basis of this type of resistance in two species, is it common to other biochemical mechanisms? Other possibilities include mutations in structural genes for the enzyme or target protein of interest, or mutations in regulatory genes which could alter the rate of transcription or processing of the structural gene (68). Indeed, mutations of structural genes are already known to confer fungicide and herbicide resistance (this volume). Is there a relationship of multiple gene families to genetic mechanisms of resistance? Multiple gene families are known for monooxygenases and glutathione transferases, two clusters of tightly linked carboxylester hydrolase genes occur on chromosome 8 of the mouse (69) and various ion channel genes have analogous construction suggesting an ancestral channel. Gene families might have arisen from duplication of an ancestral gene followed by divergence of structure through evolution. Might a similar mechanism apply to the rapid evolution of resistance? Can individual alleles for resistance be detected in the field? This will be necessary in order to develop sound resistance management strategies. An example of the complexity of the problem is analysis of methyl parathion resistance, in which we have preliminary evidence for six possible mechanisms in tobacco budworm (Table V). Many important pests have multiple mechanisms toward each group of insecticides. Some are accumulating multiple resistance at an alarming rate. How does multiple resistance arise genetically and why is it stable in the field? Are there any similarities to multiple drug resistant bacterial strains, in which transposable elements carrying several resistance genes can be transmitted among strains, and perhaps across species? Can we observe the dynamics of several r



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Figure 7


Comparison of protein ion channels. Like-shaded areas within each channel are coded by one gene. See text for details.


74



individual genes in field populations? Recent progress in this area based on biochemical techniques must continue while genetic techniques are added for improved detection of specific resistance alleles (70). Genetic Studies of Important Pest Species. Linkage mapping of insecticide resistance genes in important pests is needed for better understanding of this problem. The new power of molecular genetics provides the means to study mechanisms directly in pests of greatest interest to agriculture and public health. Linkage maps can be constructed for even those pests with many chromosomes and no prior linkage information. Extrapolation of results from only a few model species could become unnecessary. The process can be accelerated and expanded to new species by employing restriction fragment length polymorphism (rflp) mapping. We are constructing a linkage map of the tobacco budworm using both enzyme polymorphisms, which now mark fourteen of 30 autosomal linkage groups, and adding rflp markers. We have observed linkage of acetylcholinesterase resistance to another marker enzyme and are attempting to map other resistance genes. (Heckel, D. G.; Bryson, P. K.; Brown, T. M., Clemson University, personal communication, 1988). One application of the map will be to isolate mechanisms to study their potency. If structural gene mutations confer resistance, this can be confirmed, since the gene of interest should map to the same locus as the resistance trait. Genes for resistance can be isolatedfromthe pest of interest by recombinant DNA techniques. Table V. Putative Mechanisms of Methyl Parathion Resistance in Several Strains of the Tobacco Budworm, Heliothis virescens Mechanism

Laboratory (Reference)

Monooxygenase Enhanced detoxication (TCPB synergism) Less activation to oxon

This paper Konno et al. (78)

Arylester hydrolase

Konno et al. (22)

Carboxylester hydrolase (TBPT, EPN synergism)

Brown (16)

Altered acetylcholinesterase Slower inhibition reaction Enhanced recovery from phosphinate

This paper This paper

Acknowledgments Technical contribution No. 2922 of the South Carolina Agricultural Experiment Station. I thank P. K. Bryson for technical assistance, and T. Konno and W. C. Dauterman of North Carolina State University for collaboration and sharing data. Literature cited 1. 2. 3. 4.

Sawicki, R. M. Pestic. Sci. 1973, 4, 501-12. Hoyer, R. F.; Plapp, F.W., Jr. J. Econ. Entomol. 1968, 61, 1269-76. Plapp, F. W. Jr. In Pesticide Resistance Strategies and Tactics for Management; National Academy of Sciences: Washington, DC, 1986; p 74. Tsukamoto, M. In Pest Resistance to Pesticides; Georghiou, G. P.; Saito, T., Eds.; Plenum Press: New York, 1983; pp 71-98.


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6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. 32. 33. 34. 35. 36. 37. Physiol. 38. 39. 40. 41. 42. 43. 44. 45. 46.

Biochemical & Genetic Mechanisms ofInsecticide Resistance

Brown, A. W. A. In Pesticides in the Environment; White-Stevens, R., Ed.; Marcel Dekker: New York, 1971; Vol. 1, p 457. Georghiou, G. P.Exp.Parasitol. 1969, 26, 224-55. Hodgson, E.; Kulkarni, A. P. In Pest Resistance to Pesticides; Georghiou, G. P.; Saito,T., Eds.; Plenum Press: New York, 1983; pp 207-47. Raffa, K. F.; Priester, T. M. J. Agr. Entomol. 1985, 2, 27-45. Liu, M. Y.; Chen, J. S.; Sun, C. N. J. Econ. Entomol. 1984,77,851-56. Daly, J. C. Pestic. Sci. 1988, 23, 165-76. Payne, G. T. Dissertation, Clemson Univ., 1987. Dittrich, V.; Luetkemeier, N.; Voss; G. J. Econ. Entomol. 1979,72,380-84. Payne, G. T. Thesis, Clemson Univ., 1982. Knowles, C. O.; Roulston, W. J. J. Aust. Entomol. Soc. 1972,11,349-50. Hollingworth, R. M.; Murdock, L. L. Science 1980, 208, 74-76. Payne, G.T.; Brown, T.M. J. Econ. Entomol. 1984, 77, 294-97. Nebert, D. W.; Gonzalez, F. J. Ann. Rev. Biochem. 1987, 56, 945-93. Feyereisen, R.; Koener, J.; Farnsworth, D.; Nebert, D. PNAS. 1989,86,1465-69. Agosin, M. In Cytochrome P-450, Biochemistry, Biophysics, and Environmental Implications; Hietanen, E.; Laitinen, M.; Hanniner, O., Eds.; Elsevier: Amsterdam, 1982; pp 661-69. Ronis, M. J. J.; Hodgson, E.; Dauterman, W. C. Pestic. Biochem. Physiol. 1988, 32, 74-90. Yu, S. J. Pestic. Biochem. Physiol. 1983, 19, 330-36. Waters, L. C.; Nix, C. E. Pestic. Biochem. Physiol. 1988,30,214-27. Zeigler, R.; Whyard, S.; Downe, A. E. R.; Wyatt, G. R.; Walker, V. K. Pestic. Biochem. Physiol. 1988, 28, 279-85. Beeman, R. W. J. Econ. Entomol. 1983,76,737-40. Beeman, R. W.; Schmidt, B. A. J. Econ. Entomol. 1982,75,945-49. Matsumura, F.; Voss, G. J. Insect Physiol. 1965, 11, 147-60. Dowd, P. F.; Gagne, C. C.; Sparks, T. Pestic. Biochem. Physiol. 1987, 28, 9-16. Dowd, P. F.; Sparks, T. Pestic. Biochem. Physiol. 1986, 25, 73-81. Herath, P. R.; Hemingway, J; Weerasinghe, I. S.; Jayawardena, K. G. I. Pestic. Biochem. Physiol. 1987, 29, 157-62. Ashour, M-B. A.; Harshman, L. G.; Hammock, B. D. Pestic. Biochem. Physiol. 1987, 29, 97-111. Shono, T. Appl. Ent. Zool. 1983, 18, 407-15. Mouches, C.; Pasteur, N.; Berge, J.B.; Hyrien, O.; Raymond, M.; de Saint Vincent, B. R.; de Silvestri, M.; Georghiou, G. P. Science 1986, 233, 778-80. Field, L. M.; Devonshire, A. L.; Forde, B. G. Biochem. J. 1988, 251, 309-12. Devonshire, A. L. Biochem. J. 1977, 167, 675-83. Krisch, K. Biochem. Biophys. Acta 1966, 122, 265-80. Bryson, P. K.; Brown, T. M. Biochem. Pharmacol. 1985,34,1789-94. Motoyama, N.; Kao, L. R.; Lin, P. T.; Dauterman, W. C. Pestic. Biochem. 1984, 21, 139-47. Devonshire, A. L.; Moores, G. D.; Ffrench-Constant, R. H. Bull. Ent. Res. 1986, 76, 97-107. Raymond, M.; Pasteur, N.; Fournier, D.; Cuany, M.; Berge, J.; Magnin, M. C. R. Acad. Sci. III. 1985,300,509-12. Takahashi, M.; Yasutomi, K. J. Med. Entomol. 1987, 24, 595-603. Main, A. R.; Dauterman, W. C. Can. J. Biochem. 1967, 45, 757-71. Zimmerman, J. K.; Brown, T. M. J. Agric. Food Chem. 1986,34,516-20. Main,A.R. Biochem. J. 1960,74,10-20. Grothusen, J. R.; Bryson, P. K.; Zimmerman J. K.; Brown, T. M. J. Agric. Food Chem. 1986, 34, 513-15. Brealey, C. J.; Walker, C. H.; Baldwin, B. C. Pestic. Sci. 1980, 11, 546-54. Mulbry, W. W.; Karns, J. S.; Kearney, P. C.; Nelson, J. O.; McDaniel, C. S.; Wild, J. R.Appl.Environ. Microbiol. 1986, 51, 926-30.


76 47. 48. 49. 50. 51.


52. 53. 54. 55. 56. 57. 58. 59. 60. 61. 62. 63. 64. 65. 66. 67. 68. 69. 70. 71. 72. 73. 74. 75. 76. 77. 78. 79.

MANAGING RESISTANCE TO AGROCHEMICALS Dauterman, W. C. In Pest Resistance to Pesticides; Georghiou, G. P.; Saito, T., Eds.; Plenum Press: New York, 1983; pp 229-47. Grove, G.; Zarlengo, R. P.; Timmerman, K. P.;, Li, N.; Tam, M. F.; Tu, C-P. D. Nucleic Acids Res. 1988, 16, 425-38. Ottea, J. A.; Plapp, F. W. Jr. Pestic. Biochem. Physiol. 1984, 22, 203-208. Wadleigh, R.W.;Yu, S. J. Insect Biochem. 1987, 17, 759-64. Hama, H. In Pest Resistance to Pesticides; Georghiou, G. P.; Saito, T., Eds.; Plenum Press: New York, 1983; pp 299-331. Moores, G. D.; Devonshire, A. L.; Denholm, I. Bull. Ent. Res. 1988, 78, 537-44. Tripathi, R. K. Pestic. Biochem. Physiol. 1976, 6, 30-34. Yamamoto, I.; Takahashi, Y.; Kyomura, N. In Pest Resistance to Pesticides; Georghiou, G. P.; Saito, T., Eds.; Plenum Press: New York, 1983; pp 579-94. Anber, H. A. I.; Overmeer, W. P. J. Pestic. Biochem. Physiol. 1988, 31, 91-98. Grothusen, J. R.; Brown, T. M. Pestic. Biochem. Physiol. 1986, 26, 100-109. Hall, L. M. C.; Spierer, P. EMBO J., 1986,5,2949-54. Narahashi, T. In Pest Resistance to Pesticides; Georghiou, G. P.; Saito, T., Eds.; Plenum Press: New York, 1983; pp 333-352. Rossignol, D. P. Pestic. Biochem. Physiol. 1988, 32, 146-52. Kasbekar, D. P.; Hall, L. M. Pestic. Biochem. Physiol. 1988, 32, 135-45. Payne, G. T.; Blenk, R. G.; Brown, T. M. J. Econ. Entomol., 1987, 81, 65-73. Noda, M.; Ikeda, T.; Suzuki, H.; Takeshima, H.; Takahashi, T.; Kuno, M.; Numa, S. Nature 1986,322,826-28. Salkoff, L.; Butler, A.; Wei, A.; Scavarda, N.; Giffen, K.; Ifune, C.; Goodman, R.; Mandel, G. Science 1987, 237, 744-49. Schofield, P. R.; Darlison, M. G.; Fujita, N.; Burt, D. R.; Stephenson, F. A.; Rodriquez, H.; Rhee, L. M.; Ramachandran, J.; Reale, V.; Glencorse, T. A.; Seeburg, P. H.; Barnard, E. A. Nature 1987,328,221-27. Blair, L. A. C.; Levitan, E. S.; Marshall, J.; Dionne, V. E.; Barnard, E. A. Science 1988, 242, 577-79. Kadous, A. A.; Ghiasuddin, S. M.; Matsumura, F.; Scott, J. G.; Tanaka, K. Pestic. Biochem. Physiol. 1983, 19, 157-66. Imoto, K.; Methfessel, C.; Sakmann, B.; Mishina, M.; Yori, Y.; Konno, T.; Fukuda,K.; Kurasaki, M.; Bujo, H.; Fujita, Y.; Numa, S. Nature 1986, 324, 67074. Plapp, F. W. Jr. Pestic. Biochem. Physiol. 1984, 22, 194-201. Berning, W.; DeLooze, S. M.; Von Diemling, O. Comp. Biochem. Physiol. 1985, 80, 859-66. Brogdon, W.G., Hobbs, J.H., St. Jean, Y., Jacques, J.R. and Charles, L.B. J. Am. Mosquito Contr. Assoc. 1988, 4, 152-157. Kao, L. R.; Motoyama, N.; Dauterman, W. C. Pestic. Biochem. Physiol. 1985, 23, 228-39. Sultatos, L. G.; Basker, K. M.; Shao, M.; Murphy, S. D. Mol. Pharmacol. 1984, 26, 99-104. Halliday, W. R. J. Stored Prod. Res. 1988, 24, 91-99. Main, A. R.; Braid, P. E. Biochem. J. 1962,84,255-63. Lin, P. T.; Main, A. R.; Motoyama, N.; Dauterman, W. C. Pestic. Biochem. Physiol. 1984, 22, 110-16. Suzuki, T.; Miyamoto, J. Pestic. Biochem. Physiol. 1978, 8, 186-98. Abdel-Aal, Y. A.; Soderlund, D. M. Pestic. Biochem. Physiol. 1980, 14, 282-89. Wilson, T. G. J. Econ. Entomol. 1988,81,22-27. Konno, T.; Hodgson, E.; Dauterman, W. C. Pestic. Biochem. Physiol. 1989, 33, 189-99.

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